Insects and other pests cost farmers billions of dollars annually in crop losses and in the expense of keeping these pests under control. The losses caused by insect pests in agricultural production environments include decreases in crop yield, reduced crop quality, and increased harvesting costs. Insect pests are also a burden to vegetable and fruit growers, to producers of ornamental flowers, and to home gardeners and homeowners.
Cultivation methods, such as crop rotation and the application of high levels of nitrogen fertilizers, have partially addressed problems caused by agricultural pests. However, economic demands on the utilization of farmland restrict the use of crop rotation. In addition, overwintering traits of some insects are disrupting crop rotations in some areas.
Thus, synthetic chemical insecticides are relied upon most heavily to achieve a sufficient level of control. However, the use of synthetic chemical insecticides can have several drawbacks. For example, the use of some of these chemicals can adversely affect many beneficial insects. Target insects have also developed resistance to some chemical pesticides. This has been partially alleviated by various resistance management strategies, but there is an increasing need for alternative pest control agents. Furthermore, very high populations of larvae, heavy rains, and improper calibration of insecticide application equipment can result in poor control. The improper use of insecticides raises environmental concerns such as contamination of soil and of both surface and underground water supplies. Residues can also remain on treated fruits, vegetables, and on other treated plants. Working with some insecticides can also pose hazards to the persons applying them. Therefore, synthetic chemical pesticides are being increasingly scrutinized for their potential toxic environmental consequences. Stringent new restrictions on the use of pesticides and the elimination of some effective pesticides from the market place could limit economical and effective options for controlling damaging and costly pests.
Because of the problems associated with the use of synthetic chemical pesticides, there exists a clear need to limit the use of these agents and a need to identify alternative control agents. The replacement of synthetic chemical pesticides, or combination of these agents with biological pesticides, could reduce the levels of toxic chemicals in the environment.
Some biological pesticidal agents that are now being used with some success are derived from the soil microbe Bacillus thuringiensis (B.t.). The soil microbe Bacillus thuringiensis (B.t.) is a Gram-positive, spore-forming bacterium. Most strains of B.t. do not exhibit pesticidal activity. Some B.t. strains produce, and can be characterized by, parasporal crystalline protein inclusions. These inclusions often appear microscopically as distinctively shaped crystals. Some B.t. proteins are highly toxic to pests, such as insects, and are specific in their toxic activity. Certain insecticidal B.t. proteins are associated with the inclusions. These “δ-endotoxins” are different from exotoxins, which have a non-specific host range. Other species of Bacillus also produce pesticidal proteins.
Certain Bacillus toxin genes have been isolated and sequenced, and recombinant DNA-based products have been produced and approved for use. In addition, with the use of genetic engineering techniques, various approaches for delivering these toxins to agricultural environments are being perfected. These include the use of plants genetically engineered with toxin genes for insect resistance and the use of stabilized intact microbial cells as toxin delivery vehicles. Thus, isolated Bacillus toxin genes are becoming commercially valuable.
Commercial use of B.t. pesticides was initially restricted to targeting a narrow range of lepidopteran (caterpillar) pests. Preparations of the spores and crystals of B. thuringiensis subsp. kurstaki have been used for many years as commercial insecticides for lepidopteran pests. For example, B. thuringiensis var. kurstaki HD-1 produces a crystalline δ-endotoxin which is toxic to the larvae of a number of lepidopteran insects.
More recently, new subspecies of B.t. have been identified, and genes responsible for active δ-endotoxin proteins have been isolated. Höfte and Whiteley classified B.t. crystal protein genes into four major classes (Höfte, H., H. R. Whiteley [1989] Microbiological Reviews 52(2):242-255). The classes were CryI (Lepidoptera-specific), CryII (Lepidoptera- and Diptera-specific), CryIII (Coleoptera-specific), and CryIV (Diptera-specific). The discovery of strains specifically toxic to other pests has been reported. For example, CryV and CryVI were proposed to designate a class of toxin genes that are nematode-specific.
The Lepidopteran-specific CryI crystal proteins, in their natural state, are approximately 130- to 140-kDa proteins, which accumulate in bipyramidal crystalline inclusions during the sporulation of B. thuringiensis. These proteins are protoxins which solubilize in the alkaline environment of the insect midgut and are proteolytically converted by crystal-associated or larval-midgut proteases into a toxic core fragment of 60 to 70 kDa. This activation can also be carried out in vitro with a variety of proteases. The toxic domain is localized in the N-terminal half of the protoxin. This was demonstrated for CryIA(b) and CryIC proteins through N-terminal amino acid sequencing of the trypsin-activated toxin. Höfte et al. 1989. Cleavage occurs on the C-terminal end of a conserved region called “Block 5,” thus forming the C-terminus of the core toxin. A short, N-terminal protoxin segment can also be processed off. The N-terminal cleavage site is also highly conserved for CryIA and CryID proteins, suggesting that for these proteins, the N terminus of the toxic fragment is localized at the same position. CryIB, however, is different from the other CryI proteins in this region. It was not known whether this protein is also processed at the N terminus. Höfte et al. 1989.
Deletion analysis of several cryI genes further confirmed that the 3′ half of the protoxin is not required for toxic activity. One of the shortest reported toxic fragments was localized between codons 29 and 607 for CryIAb. Further removal of four codons from the 3′ end or eight codons from the 5′ end completely abolished the toxic activity of the gene product. Similar observations were made for the cryIA(a) and cryIA(c) genes. Höfte et al. 1989.
The cryII genes encode 65-kDa proteins which form cuboidal inclusions in strains of several subspecies. These crystal proteins were previously designated “P2” proteins, as opposed to the 130-kDa P1 crystal proteins present in the same strains. Höfte et al. 1989.
A cryIIA gene was cloned from B. thuringiensis subsp. kurstaki HD-263 and expressed in Bacillus megaterium. Cells producing the CryIIA protein were toxic for the lepidopteran species Heliothis virescens and Lymantria dispar as well as for larvae of the dipteran Aedes aegypti. Widner and Whitely (1989, J. Bacteriol. 171:965-974) cloned two related genes (cryIIA and cryIIB) from B. thuringiensis subsp. kurstaki HD-1. Both genes encode proteins of 633 amino acids with a predicted molecular mass of 71 kDa (slightly larger than the apparent molecular mass determined for the P2 proteins produced in B. thuringiensis). Although the CryIIA and CryIIB proteins are highly homologous (87% amino acid identity), they differ in their insecticidal spectra. CryIIA is active against both a lepidopteran (Manduca sexta) and a dipteran (Aedes aegypti) species, whereas cryIIB is toxic only to the lepidopteran insect. Höfte et al. 1989. The CryII toxins, as a group, tend to be relatively more conserved at the sequence level (>80% identical) than other groups. In contrast, there are many CryI toxins, for example, including some that are less than 60% identical.
The 1989 nomenclature and classification scheme of Höfte and Whiteley for crystal proteins was based on both the deduced amino acid sequence and the host range of the toxin. That system was adapted to cover 14 different types of toxin genes which were divided into five major classes. The 1989 nomenclature scheme became unworkable as more and more genes were discovered that encoded proteins with varying spectrums of pesticidal activity. Thus, a revised nomenclature scheme was adopted, which is based solely on amino acid identity (Crickmore et al., 1998, Microbiology and Molecular Biology Reviews 62:807-813). The mnemonic “cry” has been retained for all of the toxin genes except cytA and cytB, which remain a separate class. Roman numerals have been exchanged for Arabic numerals in the primary rank, and the parentheses in the tertiary rank have been removed. Many of the original names have been retained, with the noted exceptions, although a number have been reclassified. There are now at least 37 primary classes of Cry proteins, and two primary classes of cyt toxins. Other types of toxins, such as those of WO 98/18932 and WO 97/40162, have also been discovered from B. thuringiensis. 
There are some obstacles to the successful agricultural use of Bacillus (and other biological) pesticidal proteins. Certain insects can be refractory to the effects of Bacillus toxins. Insects such as boll weevils, black cutworm, and Helicoverpa zea, as well as adult insects of most species, heretofore have demonstrated no significant sensitivity to many B.t. δ-endotoxins.
Another potential obstacle is the development of resistance to B.t. toxins by insects. B.t. protein toxins were initially formulated as sprayable insect control agents. A more recent application of B.t. technology has been to isolate and transform plants with genes that encode these toxins. Transgenic plants subsequently produce the toxins, thereby providing insect control. See U.S. Pat. Nos. 5,380,831; 5,567,600; and 5,567,862 to Mycogen Corporation. Transgenic B.t. plants are quite efficacious, and usage is predicted to be high in some crops and areas. This has caused some concern that resistance management issues may arise more quickly than with traditional sprayable applications. While a number of insects have been selected for resistance to B.t. toxins in the laboratory, only the diamondback moth (Plutella xylostella) has demonstrated resistance in a field setting (Ferre, J. and Van Rie, J., Annu. Rev. Entomol. 47:501-533, 2002).
Resistance management strategies in B.t. transgene plant technology have become of great interest (for example, as in a natural bacterium, multiple diverse toxins can be exposed on the same plant, thereby greatly reducing the chance that an insect that might be resistant to one toxin would survive to spread the resistance). Several strategies have been suggested for preserving the ability to effectively use B. thuringiensis toxins. These strategies include high dose with refuge, and alternation with, or co-deployment of, different toxins (McGaughey et al. (1998), “B.t. Resistance Management,” Nature Biotechnol 16:144-146).
Thus, there remains a great need for developing additional genes that can be expressed in plants in order to effectively control various insects. In addition to continually trying to discover new B.t. toxins, it would be quite desirable to discover other bacterial sources (distinct from B.t.) that produce toxins that could be used in transgenic plant strategies, or that could be combined with B.t.s to produce insect-controlling transgenic plants.
The recent efforts to clone insecticidal toxin genes from the Photorhabdus/Xenorhabdus group of bacteria present potential alternatives to toxins derived from B. thuringiensis. It has been known in the art that bacteria of the genus Xenorhabdus are symbiotically associated with the Steinernema nematode. Unfortunately, as reported in a number of articles, the bacteria only had pesticidal activity when injected into insect larvae and did not exhibit biological activity when delivered orally.
It has been difficult to effectively exploit the insecticidal properties of the nematode or its bacterial symbiont. Thus, it would be quite desirable to discover proteinaceous agents from Xenorhabdus bacteria that have oral activity so that the products produced therefrom could be formulated as a sprayable insecticide, or the bacterial genes encoding said proteinaceous agents could be isolated and used in the production of transgenic plants. WO 95/00647 relates to the use of Xenorhabdus protein toxin to control insects, but it does not recognize orally active toxins. WO 98/08388 relates to orally administered pesticidal agents from Xenorhabdus. U.S. Pat. No. 6,048,838 relates to protein toxins/toxin complexes, having oral activity, obtainable from Xenorhabdus species and strains.
Photorhabdus and Xenorhabdus spp. are Gram-negative bacteria that entomopathogenically and symbiotically associate with soil nematodes. These bacteria are found in the gut of entomopathogenic nematodes that invade and kill insects. When the nematode invades an insect host, the bacteria are released into the insect haemocoel (the open circulatory system), and both the bacteria and the nematode undergo multiple rounds of replication; the insect host typically dies. These bacteria can be cultured away from their nematode hosts. For a more detailed discussion of these bacteria, see Forst and Nealson, 60 Microbiol. Rev. 1 (1996), pp. 21-43.
The genus Xenorhabdus is taxonomically defined as a member of the Family Enterobacteriaceae, although it has certain traits atypical of this family. For example, strains of this genus are typically nitrate reduction negative and catalase negative. Xenorhabdus has only recently been subdivided to create a second genus, Photorhabdus, which is comprised of the single species Photorhabdus luminescens (previously Xenorhabdus luminescens) (Boemare et al., 1993 Int. J. Syst. Bacteriol. 43, 249-255). This differentiation is based on several distinguishing characteristics easily identifiable by the skilled artisan. These differences include the following: DNA-DNA characterization studies; phenotypic presence (Photorhabdus) or absence (Xenorhabdus) of catalase activity; presence (Photorhabdus) or absence (Xenorhabdus) of bioluminescence; the Family of the nematode host in that Xenorhabdus is found in Steinernematidae and Photorhabdus is found in Heterorhabditidae); as well as comparative, cellular fatty-acid analyses (Janse et al. 1990, Lett. Appl. Microbiol. 10, 131-135; Suzuki et al. 1990, J. Gen. Appl. Microbiol., 36, 393-401). In addition, recent molecular studies focused on sequence (Rainey et al. 1995, Int. J. Syst. Bacteriol., 45, 379-381) and restriction analysis (Brunel et al., 1997, App. Environ. Micro., 63, 574-580) of 16S rRNA genes also support the separation of these two genera.
The expected traits for Xenorhabdus are the following: Gram stain negative rods, white to yellow/brown colony pigmentation, presence of inclusion bodies, absence of catalase, inability to reduce nitrate, absence of bioluminescence, ability to uptake dye from medium, positive gelatin hydrolysis, growth on Enterobacteriaceae selective media, growth temperature below 37° C., survival under anaerobic conditions, and motility.
Currently, the bacterial genus Xenorhabdus is comprised of four recognized species, Xenorhabdus nematophilus, Xenorhabdus poinarii, Xenorhabdus bovienii and Xenorhabdus beddingii (Brunel et al., 1997, App. Environ. Micro., 63, 574-580). A variety of related strains have been described in the literature (e.g., Akhurst and Boemare1988 J. Gen. Microbiol., 134, 1835-1845; Boemare et al. 1993 Int. J. Syst. Bacteriol. 43, pp. 249-255; Putz et al. 1990, Appl. Environ. Microbiol., 56, 181-186, Brunel et al., 1997, App. Environ. Micro., 63, 574-580, Rainey et al. 1995, Int. J. Syst. Bacteriol., 45, 379-381).
Xenorhabdus and Photorhabdus bacteria secrete a wide variety of substances into the culture medium; these secretions include lipases, proteases, antibiotics and lipopolysaccharides. Purification of different protease fractions has clearly demonstrated that they are not involved in the oral toxic activity of P. luminescens culture medium (which has been subsequently determined to reside with the Tc proteins only). Several of these substances have previously been implicated in insect toxicity but until recently no insecticidal genes had been cloned. However, protease purification and separation will also facilitate an examination of their putative role in, for example, inhibiting antibacterial proteins such as cecropin. R. H. ffrench-Constant and Bowen, Current Opinions in Microbiology, 1999, 12:284-288. See R. H. ffrench-Constant et al. 66 AEM No. 8, pp. 3310-3329 (August 2000), for a review of various factors involved in Photorhabdus virulence of insects.
There has been substantial progress in the cloning of genes encoding insecticidal toxins from both Photorhabdus luminescens and Xenorhabdus nematophilus. Toxin-complex encoding genes from P. luminescens were examined first. See, e.g., WO 98/08932. “Parallel” genes were more recently cloned from X. nematophilus. Morgan et al., Applied and Environmental Microbiology 2001, 67:2062-69.
Four different toxin complexes (TCs)—Tca, Tcb, Tcc and Tcd—have been identified in Photorhabdus spp. Each of these toxin complexes resolves as either a single or dimeric species on a native agarose gel but resolution on a denaturing gel reveals that each complex consists of a range of species between 25-280 kDa. The ORFs that encode the TCs from Photorhabdus, together with protease cleavage sites (vertical arrows), are illustrated in FIG. 1. See also R. H. ffrench-Constant and Bowen, 57 Cell. Mol. Life Sci. 828-833 (2000).
Genomic libraries of P. luminescens were screened with DNA probes and with monoclonal and/or polyclonal antibodies raised against the toxins. Four tc loci were cloned: tca, tcb, tcc and tcd. The tca locus is a putative operon of three open reading frames (ORFs), tcaA, tcaB, and tcaC transcribed from the same DNA strand, with a smaller terminal ORF (tcaZ) transcribed in the opposite direction. The tcc locus also is comprised of three ORFs putatively transcribed in the same direction (tccA, tccB, and tccC). The tcb locus is a single large ORF (tcbA), and the tcd locus is composed of two ORFs (tcdA and tcdB); tcbA and tcdA, each about 7.5 kb, encode large insect toxins. TcdB has some homology to TcaC. Many of these gene products were determined to be cleaved by proteases. For example, both TcbA and TcdA are cleaved into three fragments termed i, ii and iii (e.g. TcbAi, TcbAii and TcbAiii). Products of the tca and tcc ORFs are also cleaved. See FIG. 1. See also R. H. ffrench-Constant and D. J. Bowen, Current Opinions in Microbiology, 1999, 12:284-288.
Bioassays of the Tca toxin complexes revealed them to be highly toxic to first instar tomato hornworms (Manduca sexta) when given orally (LD50 of 875 ng per square centimeter of artificial diet). R. H. ffrench-Constant and Bowen 1999. Feeding was inhibited at Tca doses as low as 40 ng/cm2. Given the high predicted molecular weight of Tca, on a molar basis, P. luminescens toxins are highly active and relatively few molecules appear to be necessary to exert a toxic effect. R. H. ffrench-Constant and Bowen, Current Opinions in Microbiology, 1999, 12:284-288.
None of the four loci showed overall similarity to any sequences of known function in GenBank. Regions of sequence similarity raised some suggestion that these proteins (TcaC and TccA) may overcome insect immunity by attacking insect hemocytes. R. H. ffrench-Constant and Bowen, Current Opinions in Microbiology, 1999, 12:284-288.
TcaB, TcbA, and TcdA all show amino acid conservation (50% identity), compared with each other, immediately around their predicted protease cleavage sites. This conservation between three different TC proteins suggests that they may all be processed by the same or similar proteases. TcbA and TcdA also share ˜50% identity overall, as well as a similar predicted pattern of both carboxy- and amino-terminal cleavage. It was postulated that these proteins might thus be homologs of one another. Furthermore, the similar, large size of TcbA and TcdA, and also the fact that both toxins appear to act on the gut of the insect, may suggest similar modes of action. R. H. ffrench-Constant and Bowen, Current Opinions in Microbiology, 1999, 12:284-288.
Deletion/knock-out studies suggest that products of the tca and tcd loci account for the majority of oral toxicity to lepidopterans. Deletion of either of the tca or tcd genes greatly reduced oral activity against Manduca sexta. That is, products of the tca and tcd loci are oral lepidopteran toxins on their own; their combined effect contributed most of the secreted oral activity. R. H. ffrench-Constant and D. J. Bowen, 57 Cell. Mol. Life. Sci. 831 (2000). Interestingly, deletion of either of the tcb or tcc loci alone also reduces mortality, suggesting that there may be complex interactions among the different gene products. Thus, products of the tca locus may enhance the toxicity of tcd products. Alternatively, tcd products may modulate the toxicity of tca products and possibly other complexes. Noting that the above relates to oral activity against a single insect species, tcb or tcc loci may produce toxins that are more active against other groups of insects (or active via injection directly into the insect haemocoel—the normal route of delivery when secreted by the bacteria in vivo). R. H. ffrench-Constant and Bowen, Current Opinions in Microbiology, 1999, 12:284-288.
WO 01/11029 discloses nucleotide sequences that encode TcdA and TcbA and have base compositions that have been altered from that of the native genes to make them more similar to plant genes. Also disclosed are transgenic plants that express Toxin A and Toxin B.
Of the separate toxins isolated from Photorhabdus luminescens (W-14), those designated Toxin A and Toxin B have been the subject of focused investigation for their activity against target insect species of interest (e.g., corn rootworm). Toxin A is comprised of two different subunits. The native gene tcdA encodes protoxin TcdA. As determined by mass spectrometry, TcdA is processed by one or more proteases to provide Toxin A. More specifically, TcdA is an approximately 282.9 kDa protein (2516 aa) that is processed to provide TcdAi (the first 88 amino acids), TcdAii (the next 1849 aa; an approximately 208.2 kDa protein encoded by nucleotides 265-5811 of tcdA), and TcdAiii, an approximately 63.5 kDa (579 aa) protein (encoded by nucleotides 5812-7551 of tcdA). TcdAii and TcdAiii appear to assemble into a dimer (perhaps aided by TcdAi), and the dimers assemble into a tetramer of four dimers. Toxin B is similarly derived from TcbA.
While the exact molecular interactions of the TC proteins with each other, and their mechanism(s) of action, are not currently understood, it is known, for example, that the Tca toxin complex of Photorhabdus is toxic to Manduca sexta. In addition, some TC proteins are known to have “stand alone” insecticidal activity, while other TC proteins are known to potentiate or enhance the activity of the stand-alone toxins. It is known that the TcdA protein is active, alone, against Manduca sexta, but that TcdB and TccC, together, can be used to enhance the activity of TcdA. Waterfield, N. et al., Appl. Environ. Microbiol. 2001, 67:5017-5024. TcbA (there is only one Tcb protein) is another stand-alone toxin from Photorhabdus. The activity of this toxin (TcbA) can also be enhanced by TcdB together with TccC-like proteins.
U.S. Patent Application 20020078478 provides nucleotide sequences for two potentiator genes, tcdB2 and tccC2, from the tcd genomic region of Photorhabdus luminescens W-14. It is shown therein that coexpression of tcdB and tccC1 with tcdA results in enhanced levels of oral insect toxicity compared to that obtained when tcdA is expressed alone. Coexpression of tcdB and tccC1 with tcdA or tcbA provide enhanced oral insect activity.
As indicated in the chart below, TccA has some level of homology with the N terminus of TcdA, and TccB has some level of homology with the C terminus of TcdA. TccA and TccB are much less active on certain test insects than is TcdA. TccA and TccB from Photorhabdus strain W-14 are called “Toxin D.” “Toxin A” (TcdA), “Toxin B” (TcbA), and “Toxin C” (TcaA and TcaB) are also indicated below. Furthermore, TcaA has some level of homology with TccA and likewise with the N terminus of TcdA. Still further, TcaB has some level of homology with TccB and likewise with the N terminus of TcdA. TccA and TcaA are of a similar size, as are TccB and TcaB. TcdB has a significant level of similarity (both in sequence and size) to TcaC.
Photorhabdusstrain W14Some homologyPhotorhabdusnomenclatureto:TcaAToxin CTccATcaBTccBTcaCTcdBTcbAToxin BTccAToxin DTcdA N terminusTccBTcdA C terminusTccCTcdAToxin ATccA + TccBTcdBTcaC
The insect midgut epithelium contains both columnar (structural) and goblet (secretory) cells. Ingestion of tca products by M. sexta leads to apical swelling and blebbing of large cytoplasmic vesicles by the columnar cells, leading to the eventual extrusion of cell nuclei in vesicles into the gut lumen. Goblet cells are also apparently affected in the same fashion. Products of tca act on the insect midgut following either oral delivery or injection. R. H. ffrench-Constant and D. J. Bowen, Current Opinions in Microbiology, 1999, 12:284-288. Purified tca products have shown oral toxicity against Manduca sexta (LD50 of 875 ng/cm2). R. H. ffrench-Constant and D. J. Bowen, 57 Cell. Mol. Life Sci. 828-833 (2000).
WO 99/42589 and U.S. Pat. No. 6,281,413 disclose TC-like ORFs from Photorhabdus luminescens. WO 00/30453 and WO 00/42855 disclose TC-like proteins from Xenorhabdus. WO 99/03328 and WO 99/54472 (and U.S. Pat. Nos. 6,174,860 and 6,277,823) relate to other toxins from Xenorhabdus and Photorhabdus. 
Relatively recent cloning efforts in Xenorhabdus nematophilus also appear to have identified novel insecticidal toxin genes with homology to the P. luminescens tc loci. See, e.g., WO 98/08388 and Morgan et al., Applied and Environmental Microbiology 2001, 67:2062-69. In R. H. ffrench-Constant and D. J. Bowen, Current Opinions in Microbiology, 1999, 12:284-288, cosmid clones were screened directly for oral toxicity to another lepidopteran, Pieris brassicae. One orally toxic cosmid clone was sequenced. Analysis of the sequence in that cosmid suggested that there are five different ORF's with similarity to Photorhabdus tc genes; orf2 and orf5 both have some level of sequence relatedness to both tcbA and tcdA, whereas orf1 is similar to tccB, orf3 is similar to tccC and orf4 is similar to tcaC. Importantly, a number of these predicted ORFs also share the putative cleavage site documented in P. luminescens, suggesting that active toxins may also be protealytically processed.
There are five typical Xenorhabdus TC proteins: XptA1, XptA2, XptB1, XptC1, and XptD1. XptA1 is a “stand-alone” toxin. XptA2 is another TC protein from Xenorhabdus that has stand-alone toxin activity. See GENBANK Accession No. AJ308438 for sequences from Xenorhabdus nematophilus. XptB1 and XptC1 are the Xenorhabdus potentiators that can enhance the activity of either (or both) of the XptA toxins. XptD1 has some level of homology with TccB. XptC1 has some level of similarity to TcaC. The XptA2 protein of Xenorhabdus has some degree of similarity to the TcdA protein. XptB1 has some level of similarity to TccC.
The finding of somewhat similar, toxin-encoding loci in these two different bacteria is interesting in terms of the possible origins of these virulence genes. The X. nematophilus cosmid also appears to contain transposase-like sequences whose presence may suggest that these loci can be transferred horizontally between different strains or species of bacteria. A range of such transfer events may also explain the apparently different genomic organization of the tc operons in the two different bacteria. Further, only a subset of X. nematophilus and P. luminescens strains appear toxic to M. sexta, suggesting either that different strains lack the tc genes or that they carry a different tc gene compliment. Detailed analysis of both a strain and toxin phylogeny within, and between, these bacterial species should help clarify the likely origin of the toxin genes and how they are maintained in different bacterial populations. R. H. ffrench-Constant and Bowen, Current Opinions in Microbiology, 1999, 12:284-288.
TC proteins and genes have more recently been described from other insect-associated bacteria such as Serratia entomophila, an insect pathogen. Waterfield et al., TRENDS in Microbiology, Vol. 9, No. 4, April 2001.
In summary, toxin complex proteins from P. luminescens and X. nematophilus appear to have little homology to previously identified bacterial toxins and should provide useful alternatives to toxins derived from B. thuringiensis. Although they have similar toxic effects on the insect midgut to other orally active toxins, their precise mode of action remains obscure. Future work could clarity their mechanism of action.
Although some Xenorhabdus TC proteins were found to “correspond” (have a similar function and some level of sequence homology) to some of the Photorhabdus TC proteins, a given Photorhabdus protein shares only about 40% sequence identity with the “corresponding” Xenorhabdus protein. This is illustrated below for four “stand-alone” toxins:
Identity to P.l. W-14 TcbAIdentity to P.l. W-14 TcdAXwi XptA144%46%Xwi XptA241%41%(For a more complete review, see, e.g., Morgan et al., “Sequence Analysis of Insecticidal Genes from Xenorhabdus nematophiles PMFI296,” Vol. 67, Applied and Environmental Microbiology, May 2001, pp. 2062-2069.)
Bacteria of the genus Paenibacillus are distinguishable from other bacteria by distinctive rRNA and phenotypic characteristics (C. Ash et al. (1993), “Molecular identification of rRNA group 3 bacilli (Ash, Farrow, Wallbanks and Collins) using a PCR probe test: Proposal for the creation of a new genus Paenibacillus,” Antonie Van Leeuwenhoek 64:253-260). Comparative 16S rRNA sequence analysis demonstrated that the genus Bacillus consisted of at least five phyletic lines. Ribosomal RNA group 3 bacilli (of Ash, Farrow, Wallbanks, and Collins (1991), comprising Bacillus polymyxa and close relatives), is phylogenetically so removed from Bacillus subtilis (the type species of the genus and other aerobic, endospore-forming bacilli) that they were reclassified as a new genus, Paenibacillus. 
Some species in this genus were known to be pathogenic to honeybees (Paenibacillus Larvae) and scarab beetle grubs (P. popilliae and P. lentimorbus). Some other Paenibacillus species that have been found to be associated with honeybees, but they are non-pathogens. At least 18 additional species are known in this genus, including P. thiaminolyticus; they have no known insect association (Shida et al., 1997; Pettersson et al., 1999). Scarabs (coleopterans) are serious pests of turf nurseries, and food crops throughout North America, and are of quarantine concern. See U.S. Department of Agriculture, Agricultural Research Service website.
P. larvae, P. popilliae, and P. lentimorbus are considered obligate insect pathogens involved with milky disease of scarab beetles (D. P. Stahly et al. (1992), “The genus Bacillus: insect pathogens,” p. 1697-1745, In A. Balows et al., ed., The Procaryotes, 2nd Ed., Vol. 2, Springer-Verlag, New York, N.Y.). These three Paenibacillus species are characteristically slow-growing, fastidious organisms that cause disease by an invasive process in which the bacteria cross the midgut and proliferate to high numbers in the hemolymph and other tissues. For all three species, some general indications of protein involvement in insect pathogenicity have been proposed; however, no specific role for a specific protein has been demonstrated. Stahly et al. concluded for P. larvae that a question of the involvement of a toxin is an open one, and that the precise cause of death in milky disease (of beetles) is not understood.
A beetle (coleopteran) toxin, Cry18, has been identified in strains of P. popilliae and P. lentimorbus. Cry18 has about 40% identity to Cry2 proteins (Zhang et al., 1997; Harrison et al., 2000). While Zhang et al. (1997) speculate that Cry18 attacks the midgut to facilitate entry of vegetative cells to the hemocoel, Harrison et al. note that there is no direct evidence for this role and further state that “the role, if any, of the paraspore protein in milky disease is unknown.” J. Zhang et al. (1997), “Cloning and Analysis of the First cry Gene from Bacillus popilliae,” J. Bacteriol. 179:4336-4341; H. Harrison et al. (2000), “Paenibacillus Associated with Milky Disease in Central and South American Scarabs,” J. Invertebr. Pathol. 76(3):169-175.
Stahly et al., Zhang et al., and Harrison et al. all point to the contrast in evidence for the role of crystal proteins of B. thuringiensis in intoxication of insects (where the high frequency of insect symptoms can be explained by the properties of the specific crystal proteins), versus the case of Paenibacillus and milky disease (where there is no such tie to the effects of a specific toxin).
Thus, while some species of Paenibacillus were known to be pathogenic to certain coleopterans and some associated with honeybees, no strain of Paenibacillus was heretofore known to be toxic to lepidopterans. Likewise, TC proteins and lepidopteran-toxic Cry proteins have never been reported in Paenibacillus. 